Spinning-Disk Confocal Microscopy of Yeast

C H A P T E R

T W E N T Y- T H R E E

Spinning-Disk Confocal Microscopy of Yeast Kurt Thorn Contents 582 583 585 585 585 586 590 593 594 594 594 594 597 598 600 600

1. Introduction 2. Building a Spinning-Disk Confocal Microscope 2.1. Microscope base 2.2. Scanhead 2.3. Lasers and filters 2.4. Choice of objective 2.5. Cameras 2.6. Other hardware considerations 2.7. Software 2.8. System integration 3. Sample Preparation 3.1. Fluorescent tagging and choice of fluorescent protein 3.2. Minimizing autofluorescence 3.3. Mounting Acknowledgments References

Abstract Spinning-disk confocal microscopy is an imaging technique that combines the out-of-focus light rejection of confocal microscopy with the high sensitivity of wide-field microscopy. Because of its unique features, it is well suited to highresolution imaging of yeast and other small cells. Elimination of out-of-focus light significantly improves the image contrast and signal-to-noise ratio, making it easier to resolve and quantitate small, dim structures in the cell. These features make spinning-disk confocal microscopy an excellent technique for studying protein localization and dynamics in yeast. In this review, I describe the rationale behind using spinning-disk confocal imaging for yeast, hardware considerations when assembling a spinning-disk confocal scope, and methods

Department of Biochemistry and Biophysics, University of California at San Francisco, San Francisco, California, USA Methods in Enzymology, Volume 470 ISSN 0076-6879, DOI: 10.1016/S0076-6879(10)70023-9

#

2010 Elsevier Inc. All rights reserved.

581

582

Kurt Thorn

for strain preparation and imaging. In particular, I discuss choices of objective lens and camera, choice of fluorescent proteins for tagging yeast genes, and methods for sample preparation.

1. Introduction In recent years, fluorescent protein tagging combined with microscopy has become a powerful tool for imaging subcellular localization in live yeast cells (Davis, 2004; Kohlwein, 2000; Rines et al., 2002). A particularly popular imaging method is spinning-disk confocal microscopy, which provides a way around one of the fundamental limitations of fluorescence microscopy: the objective lens captures light not only from the region of the sample that is in focus but also from regions in the sample above or below the focus plane. This out-of-focus light reduces contrast in the image and obscures in-focus information. Spinning-disk confocal microscopy has become such a powerful tool for imaging yeast because it provides much better optical sectioning than a conventional fluorescence microscope, but much better sensitivity that a laser-scanning confocal. In general, laser-scanning confocal microscopes lack the sensitivity required to image small dim objects, and are unlikely to produce good images of any yeast sample that is not very bright. Conversely, conventional fluorescence microscopes can be very sensitive but do not reject out-of-focus light and so cannot acquire optical sections or produce high-resolution 3D reconstructions of the cell. Spinning-disk confocals can acquire high-resolution optical sections with good sensitivity and thus fill a niche between conventional fluorescence microscopes and laser-scanning confocals. They are particularly good when imaging small dynamic processes in vivo, such as cytoskeletal dynamics or vesicle and organelle movement. For imaging requiring less resolution, such as looking at translocation between the cytoplasm and nucleus or the abundance of a transcriptional reporter, they have relatively little advantage over a conventional fluorescence microscope. There are two general ways to deal with out-of-focus light; it can either be blocked from reaching the detector, as in confocal microscopy, or it can be computationally removed after the fact, as in deconvolution microscopy (Shaw, 2006). In confocal microscopy, a pinhole in the excitation light path excites a single spot in the sample. A confocal pinhole in the detection path then blocks all light that does not originate from the excited spot. Traditional laser-scanning confocal microscopes suffer from two drawbacks for live cell work. First, because they require scanning a single illumination spot over the sample, they are typically slow, acquiring approximately 1 frame per second (fps). Second, and more importantly, due to the detectors used and

Spinning-Disk Confocal Microscopy of Yeast

583

to the very short integration times at each pixel in the image, they are relatively insensitive. Spinning-disk confocal microscopes (reviewed in Toomre and Pawley, 2006) avoid both of these problems by replacing the single pinhole in a laserscanning confocal with multiple pinholes that simultaneously illuminate many points in the sample. These pinholes are arranged in a spiral pattern on a disk so that as the disk rotates, the pinholes sweep over every point in the sample, illuminating it uniformly. This greatly improves the speed of the spinning-disk confocal (imaging at 30 fps is routine) and, as the confocal now illuminates the entire field of view in a short period of time (typically 33 ms), it forms an image that can be recorded on a highly sensitive CCD camera (as opposed to an insensitive photomultiplier tube, as in a laser-scanning confocal). The net result is that spinning-disk confocal microscopy systems are both faster and more sensitive than laser-scanning confocals. The increase in sensitivity is striking; a spinning-disk confocal system can collect as many as 50 times more photons for a given exposure than a conventional laser-scanning confocal (Murray et al., 2007). Other confocal techniques, such as slit or line scanners, share some of the same advantages as spinning-disk confocal but have not been as extensively validated and so will not be discussed further. One common misconception about confocal microscopes, including spinning-disk confocals, is that a confocal system has higher resolution than a nonconfocal system. In theory, this is not true, in that a point-like object will be imaged to a blurry disk of the same width in both systems. That is, the width of the point spread function of both a confocal and a nonconfocal system will be the same. In practice, however, the achievable resolution of a microscope is often lower than theoretically expected due to a low signal-to-noise ratio of the image. Low signal-to-noise ratios can arise from weakly fluorescent samples, but out-of-focus light can also reduce the signal-to-noise ratio even on bright samples. Out-of-focus light contributes background to the image that reduces contrast and adds additional noise, which reduces the overall signal-to-noise ratio of the image. Thus, while the spinning-disk confocal does not improve the diffraction-limited resolution of the microscope, it may improve the achievable resolution in real samples if this is limited by out-of-focus light contributing background fluorescence and noise to the images.

2. Building a Spinning-Disk Confocal Microscope A photograph of the spinning-disk confocal in the Nikon Imaging Center at UCSF, with components labeled, is shown in Fig. 23.1A. A schematic of the major components in the optical path is shown in Fig. 23.1B.

584

Kurt Thorn

A

Emission filter wheel

EMCCD camera

CSU22 Laser fiber

B

Sample Microlens disk

Pinhole disk

Objective lens

Fiber input

Dichroic Scanhead Camera Projection lens

Emission filter wheel

Figure 23.1 Overview of a spinning-disk confocal system. (A) Photograph of the spinning-disk confocal system in the Nikon Imaging Center at UCSF. It consists of a Nikon TE2000 microscope with a temperature controlled chamber from In Vivo Biosciences. The Yokogawa CSU-22 spinning-disk scanhead is labeled, along with the optical fiber bringing excitation light to the scanhead from the laser launch. A Sutter filter wheel holding emission filters is coupled to the exit of the scanhead; a projection lens is mounted after the filter wheel. The Cascade II EMCCD camera (Photometrics) is labeled as well. (B) Schematic diagram of the spinning-disk optical path. Solid lines show the excitation path, and the short dashed lines show the emission path. Elements within the dashed box are contained within the spinning-disk confocal scanhead.

Spinning-Disk Confocal Microscopy of Yeast

585

2.1. Microscope base Spinning-disk confocal systems can be built on microscopes from any of the major manufacturers, and on either an upright or inverted system, although mounting components is somewhat simpler on an inverted scope. Microscopes from all of the major manufacturers (Leica, Nikon, Olympus, and Zeiss) have excellent optical performance so the major considerations in choosing one are software support (not all software packages support all microscopes) and the availability of other features on the microscope. Most of the major microscope manufacturers have recently introduced hardware autofocus systems on their microscopes, such as the Nikon Perfect Focus System, the Zeiss Definite Focus, or the Olympus Zero Drift. These are devices that measure the position of the sample coverslip and adjust the focus to compensate for any movement of the sample, thereby maintaining focus at all times. These autofocus systems greatly improve focal stability and are considerably more precise and faster than image-based autofocus techniques. They also greatly simplify the acquisition of time-lapse movies, and if you will be acquiring time-lapse data for more than approximately 30 min, are well worth the added expense.

2.2. Scanhead While a number of disk-scanning confocal systems exist, spinning-disk confocals from the Yokogawa Electric Corp. have come to dominate the market because they include microlenses that focus the laser light through the pinhole disk, dramatically increasing the excitation efficiency of the system (Toomre and Pawley, 2006). Other spinning-disk confocal systems do not include these microlenses, making them much less bright at equivalent laser powers. There are two different versions of the Yokogawa spinning-disk confocal scanhead: the older CSU10 and the newer CSU-X1. The CSU-X1 has several improvements over the CSU10, including a redesigned excitation path that increases the amount of excitation light reaching the sample and a higher disk rotation rate that allows for frame rates faster than 30 fps. The CSU-X1 also has an optional bypass path that allows bypassing the spinning disk for conventional wide-field imaging, and an optional motorized filter changer to allow automated filter switching in the scanhead. This is helpful if you intend to build a system with a large number of laser lines.

2.3. Lasers and filters Typically, the vendor that provides the spinning-disk scanhead will also provide the lasers and laser launch for the system. The laser launch contains the optics to combine the beams from multiple lasers together and to launch

586

Kurt Thorn

them into the single mode optical fiber attached to the spinning-disk scanhead. It will also typically have an acousto-optic tunable filter (AOTF) for rapidly switching laser lines. For spinning-disk confocal microscopy, it is probably best to avoid gas lasers (with the possible exception of Ar-ion lasers). The latest solid state lasers typically have sufficient power for confocal microscopy and are smaller, produce less heat, and have longer lifetimes than gas lasers. Typically, a 50-mW laser will supply sufficient power for spinning-disk applications, although some applications (particularly rapid imaging) may benefit from increased laser power. While the exact laser wavelengths used will depend on which fluorescent tags you wish to image, commonly used wavelengths include 405 nm (DAPI), 440 nm (CFP), 473 nm, 491 nm (GFP), 532 nm, 561 nm (RFP), and 640 nm (Cy5). You will need a dichroic beamsplitter(s) in the scanhead that matches the laser lines you are using. The dichroic beamsplitter separates the emission light from the excitation light, allowing the emitted light to be imaged onto the camera. Additionally, for multicolor imaging, you will want a filter wheel at the exit of the scanhead with emission filters that define the wavelength band you wish to detect for each channel (Fig. 23.1B). This can either be a third-party filter wheel or a CSUX optional add-on filter wheel. Alternatively, if you are using a small number of wavelengths this can be replaced with a single multipass emission filter but this may result in cross talk between different channels. Multipass emission filters are, however, useful for rapid multicolor imaging by changing the excitation wavelength as they eliminate the need for moving any mechanical parts to change wavelength. Filters with very high transmission in the passband (>90%) are now available from the major vendors (Chroma Technology, Omega Optical, and Semrock) and are highly desirable to maximize the amount of light detected by the camera.

2.4. Choice of objective Typically, for spinning-disk confocal microscopy of yeast cells one wishes to maximize the resolution of the images acquired. Ultimately, the achievable resolution is limited by the numerical aperture (NA) of the objective lens used to image the sample; the larger the numerical aperture, the higher the achievable resolution. The standard measure of resolution is given by the Rayleigh criterion, which measures the minimum distance two point sources must be separated by to be distinguished, and is given by rmin ¼ 0.61 l/NAobj, where l is the wavelength of light emitted by the sample (Inoue, 2006). Maximizing resolution, therefore, suggests the use of oil-immersion objectives with a numerical aperture of 1.4 or higher. Plan Apo objectives (corrected for both field flatness and chromatic aberration at multiple wavelengths; Keller, 2006) are available from all major manufacturers with a

Spinning-Disk Confocal Microscopy of Yeast

587

numerical aperture of 1.4. An objective with a numerical aperture of 1.4 will have a resolution limit of 220 nm for GFP. Such objectives are also well matched to the pinholes in the spinningdisk confocal. Because an objective has a finite resolution, the image of a point object smaller than the diffraction limit of the objective is blurred into a disk (the Airy disk) of radius rmin ¼ 0.61l/NAobj. For optimum confocality, the diameter of the confocal pinhole should match the diameter of this Airy disk times the magnification of the objective. When these two diameters are equal, the confocal pinhole will pass all of the in-focus light while rejecting the maximum amount of out-of-focus light. If the pinhole diameter is smaller than the Airy disk diameter, in-focus light will be blocked by the pinhole, reducing the overall efficiency of the system. If the pinhole is larger than the Airy disk, additional out-of-focus light will pass through the pinhole, reducing the confocality and contrast of the system. The Yokogawa CSU spinning-disk confocal has pinholes of diameter 50 mm, which are well matched to the Airy disk diameter of 44 mm produced by a 100/1.4 NA objective imaging GFP. One drawback of using oil-immersion objectives for imaging yeast is that using an oil-immersion objective to image into a yeast cell in an aqueous environment introduces spherical aberration due to the refractive index difference between the immersion oil and the aqueous medium. An oil-immersion objective is designed to produce aberration-free images immediately adjacent to the coverslip. As one images deeper into the sample, the thickness of the immersion oil layer will shrink, and the thickness of the water layer will increase, resulting in a change in the optical properties of the system which gives rise to spherical aberration (Fig. 23.2A). When viewed through a wide-field system (e.g., the eyepieces) spherical aberration manifests as an axial asymmetry in the point spread function, and can be recognized as haloing around point-like objects on one side of focus, with no haloing on the other side of focus (Fig. 23.2B). Spherical aberration results in the detected light from the sample no longer coming to a tight focus which reduces contrast and intensity as the broader focal spot of the spherically aberrated light from deeper in the sample will be partially cut off by the spinning-disk pinholes. This effect can be quite noticeable even for objects as thin as a yeast cell. Indeed, measurable intensity falloff can be seen after imaging as little as 2 mm into the sample (Fig. 23.2C). This spherical aberration can be eliminated in one of two ways. First, using a water-immersion objective instead of an oil-immersion objective will eliminate the change in spherical aberration with imaging depth as now the oil layer is replaced with a water layer having the same refractive index as the specimen, so that as the optical properties of the system no longer change as you image deeper into the specimen. A water-immersion objective will have a lower numerical aperture than an oil objective due to the

588

Kurt Thorn

A

Objective Immersion oil

n1

Cover glass Aqueous sample

n2

Focus into sample

Focus at coverslip

Spherical aberration

C

B

3.5 Plan Apo 100x/1.4

3

7 mm

2.5 mm

Intensity (AU)

2.5 Apo TIRF 100x/1.49 correction collar

2 1.5 1 0.5 0

Unaberrated

0

2

4

6 8 10 Z-distance ( mm)

12

14

Spherically aberrated

Figure 23.2 Spherical aberration. (A) Oil-immersion objectives are designed to focus at the coverslip (redrawn from a figure by Mats Gustafsson). Under these conditions, shown in the left-hand portion of the figure, all light rays entering the sample focus to a single point. When imaging into an aqueous sample, as shown on the right, refraction occurs at the boundary between the coverslip and the solution. This additional refraction differs for rays entering at different angles, and as a result the light is no longer tightly focused to a single spot. This is spherical aberration. (B) Unaberrated and aberrated images of beads, acquired on an epifluorescence microscope. Spherical aberration can be recognized by haloing around point objects on one side of focus. On the left is shown a series of Z slices of a 100 nm bead, taken at 0.5 mm intervals.

Spinning-Disk Confocal Microscopy of Yeast

589

lower refractive index of the immersion medium which will result in lower resolution for the objective. However, as the spherical aberration introduced by using an oil objective also reduces resolution, the unaberrated water-immersion objective may have higher resolution than the aberrated oil objective. As the sample gets thicker, the benefit from using the waterimmersion objective instead of the oil-immersion objective increases, as the induced spherical aberration increases with the distance from the coverslip. An alternate approach to eliminate spherical aberration is to add additional optics that correct for the spherical aberration. In principle, this can be done with adaptive optics approaches, but this remains highly challenging (Booth, 2007). A more practical approach is to introduce additional lenses that move as the objective moves in Z to compensate for the induced spherical aberration. One such device is the MID/SAC spherical aberration corrector available from Intelligent Imaging Innovations, which has been used to great effect for imaging samples up to 80 mm thick with a spinningdisk confocal microscope (McAllister et al., 2008). For imaging thin yeast samples, however, such an approach is probably overkill. An alternative approach is to use an objective with a correction collar, which allows the optics inside the objective to be adjusted to eliminate spherical aberration at any single focal plane. While such a correction collar (unless continuously adjusted) cannot correct for spherical aberration at all focal planes in the sample, setting it to minimize spherical aberration in the middle of the yeast cell minimizes spherical aberration over the thickness of the cell and has been found to give improved quantitation of fluorescent intensities over the full thickness of the cell (Fig. 23.2C). Oil-immersion objectives with correction collars are typically designed for total internal reflection fluorescence microscopy and so additionally benefit from having a slightly higher numerical aperture (up to 1.49). For applications where accurate three-dimensional These were taken under conditions of minimal spherical aberration, and the image of the bead (the point spread function) is relatively symmetric above and below focus. On the right is an image of the same bead with spherical aberration induced; the Z slices are now taken at 1 mm intervals. The pronounced asymmetry between focusing above and below the bead is now apparent. Observing small structures in a sample and adjusting the correction collar to minimize the asymmetry in the point spread function above and below focus is a good way to minimize spherical aberration in your images. (C) The effect of spherical aberration on quantitative intensity measurements with a spinningdisk confocal (data provided by Susanne Rafelski). A 2-mm bead was imaged with both a Plan Apo 100/1.4 NA objective (without a correction collar) and an Apo TIRF 100/1.49 NA objective, with the correction collar set to minimize spherical aberration. The average intensity from a small region in the center of the bead is plotted as a function of Z. The intensity falls off rapidly when imaging into the sample with the Plan Apo objective, but is much more symmetric when using the TIRF objective with correction collar. The image of the bead acquired with the correction collar adjusted to minimize spherical aberration better matches the actual brightness of the bead, which should be uniform throughout its thickness.

590

Kurt Thorn

intensity measurements are important, an oil- or water-immersion objective with a correction collar is highly recommended. Setting the correction collar is done by adjusting it to achieve symmetry in the point spread function above and below focus around the middle of the cell. This is done by observing the point spread function of a small object in the cell through the eyepieces while adjusting the correction collar. One of the drawbacks to the use of objectives with correction collars is that setting the collar accurately takes practice, and if the collar is set inaccurately it will introduce additional spherical aberration. For this reason, we have both a 100/1.4 NA oil lens (without a correction collar) and a 100/1.49 NA oil lens (with a correction collar) on our spinning-disk confocal. The 1.4 NA lens is used for routine applications and the 1.49 NA lens is used for more demanding 3D reconstructions. For thicker specimens, such as yeast biofilms, a water-immersion lens may give more satisfactory performance. While the specific objectives mentioned here are Nikon objectives, other microscope manufacturers have generally similar objectives. For applications where high resolution and high contrast are critical, it is probably best to compare both oil- and water-immersion objectives. The achievable resolution in a spinning-disk confocal experiment depends on the numerical aperture, magnification, and aberrations of your microscope in a complex way, and the solution with the best performance is not always predictable a priori. In particular, while water-immersion objectives have lower magnification and numerical aperture, and would therefore be expected to have lower resolution, they also have lower spherical aberration, which reduces contrast and degrades resolution. In a real experiment, the achievable resolution depends not only on the theoretical resolution of the objective but also on the intensity of the signal and the background, and on any aberrations, and so can only be determined empirically.

2.5. Cameras For any spinning-disk confocal system, you will need a camera to acquire images for analysis and publication. The camera should be highly sensitive, so as to capture as many photons as possible that arrive at it, and low noise, so as to contribute as little extra noise to the image as possible. Camera sensitivity is measured by the quantum efficiency of the camera, which measures the fraction of photons incident on the CCD that are recorded. For spinning-disk confocal microscopy, particularly on dim specimens, cameras with very high quantum efficiencies are desirable, so that nearly every photon collected by the microscope is recorded on the camera. The highest possible quantum efficiencies are achieved by back-thinning of the CCD chip, whereby the substrate the chip is grown on is physically ground away and the CCD is illuminated from the back. Illuminating the CCD chip from the back eliminates absorption and scattering by the electronics

Spinning-Disk Confocal Microscopy of Yeast

591

fabricated on the front face of the CCD and allows quantum efficiencies in the visible wavelength range of  90%. By contrast, nonback-thinned CCDs (front-illuminated CCDs) have typical quantum efficiencies of 60% (Art, 2006; Janesick, 2001). Noise in the image comes from a number of sources, of which two are dominant: photon shot noise and camera read noise. Photon shot noise occurs because photons are collected in integer numbers (so that while a given pixel may record 10 photons on average, sometimes it will record more or less) and is unavoidable. The standard deviation due to the photon shot noise is given by the square root of the number of photons collected, and so the signal-to-noise ratio is given by one over the square root of the number of photons collected. Thus, the only way to reduce the photon shot noise is by collecting more photons, for example, by exposing longer. Camera read noise is noise introduced by the digitization process in the camera. This introduced noise is proportional to the readout speed of the camera—the faster the camera is read out, the higher the read noise. A typical camera that can be readout at 10 fps, such as the Coolsnap HQ2 (Photometrics) or Orca-R2 (Hamamatsu) has 6–8 e of read noise. Very low read noise can be achieved with a very slow readout speed. For instance, the Orca-II-ER (Hamamatsu) has a read noise of 4 e, but takes 1.2 s to read out a single image. For many applications of spinning-disk confocal microscopy, such a frame rate is prohibitively slow. To achieve a fast frame rate while maintaining low noise, the solution has been to amplify the signal before reading it out. Since the signal-to-noise ratio depends on both the signal strength and the amount of noise, the signal-to-noise ratio can be increased either by amplifying the signal or by reducing the noise. Either amplifying the signal or reducing the noise by the same amount results in the same increase in the signal-to-noise ratio. Therefore, amplification can be thought of as reducing the read noise by the same factor, which is why amplified cameras often quote an effective ‘‘read noise’’ of <1 e. This was first done by using an image intensifier attached to the CCD, a so-called intensified CCD or ICCD. The image intensifier greatly amplifies the number of photons prior to their arrival at the CCD (by up to a million-fold) rendering even the high read noise typical of a fast camera negligible. However, ICCDs suffer from a number of drawbacks including low quantum efficiency and potential damage due to brief exposure to bright light, so they have been largely supplanted for routine imaging by a newer technology, the electron multiplying CCD (EMCCD). An EMCCD is essentially a normal CCD with a gain register located prior to the readout electronics. The photoelectrons from the CCD are shifted through the wells of this register at higher than normal voltages, resulting in a gain of  1% for each transfer step. That is, after a single shift event through this register, 100 electrons will on average be amplified to

592

Kurt Thorn

101 electrons. The gain register consists of several hundred to a thousand wells, so shifting through the entire register results in very high gains, up to 1000-fold (Art, 2006; Janesick, 2001). This method of amplifying the signal has several advantages over that of an ICCD. First, because the gain is located after the light-sensitive CCD, rather than in front of it like an image intensifier, the high quantum efficiency (>90%) of a back-thinned CCD can be realized. Second, the amplification process introduces remarkably little extra noise. The gain register does introduce additional shot noise, however, as the amplification process is random. As this gain noise has the same statistical properties as the photon shot noise, the resulting shot noise in the signal is increased, as if fewer photons had been collected. The net effect of this is to make it appear as if half as many photons had been gathered, so that the camera’s quantum efficiency appears to be halved. Imaging quickly necessitates short exposure times (hence few photons) and cameras with fast readout (hence high read noise). Under these conditions, noise in the image will be dominated by the camera read noise, and amplification, which reduces the effective read noise, will substantially improve the final image quality. This is often the regime that spinningdisk confocal microscopy experiments fall in, as a typical experiment involves acquiring rapid Z-stacks in time-lapse, often necessitating exposures of 100 ms or less. For this reason, most spinning-disk confocal systems are now paired with EMCCD cameras. However, if you are imaging bright samples and do not need to image quickly, it may also be worth considering conventional CCD cameras as well. EMCCDs are made by three major manufacturers: Roper Scientific, Andor Technologies, and Hamamatsu. The cameras from all three companies tend to be similar, as they all utilize the same EMCCD chips. The most popular EMCCDs (e.g., Photometrics Evolve, Andor Ixon, and Hamamatsu ImagEM) use a 512  512 pixel back-thinned EMCCD chip from e2v, which can be read out at video rate (30 fps). There is also a 1k  1k pixel back-thinned EMCCD available, but it can be only read out at 8 fps and it is substantially more expensive than the 512  512 pixel EMCCDs. There is also a front illuminated EMCCD available, but this should probably be avoided due to its substantially lower quantum efficiency compared to the back-thinned EMCCDs. To achieve optimal resolution of a microscopy system, the magnification of the image on the camera must be chosen to match the resolution limit of the objective. Specifically, for the camera to be able to accurately resolve an object of a given size, that object must span at least two pixels on the camera; this is known as Nyquist–Shannon sampling (Pawley, 2006). Therefore, for your camera to achieve the full resolution that your objective is capable of, you need to magnify the image on the camera such that one camera pixel covers a distance of less than half the resolution limit of the objective when referred to the object plane. For example, a 100/1.4 NA objective has a

Spinning-Disk Confocal Microscopy of Yeast

593

resolution limit of 220 nm for GFP, so one pixel should span < 110 nm, referred to the object plane, to achieve maximum resolution. For a camera with 16 mm pixels, one pixel will correspond to a size of 160 nm in the object plane with no magnification beyond that of the 100 objective. Therefore, to achieve maximal resolution, additional magnification must be introduced; this is most easily done by replacing the projection lens of the confocal scanhead with a lens of longer focal length. It is often useful to include additional magnification beyond that required by Nyquist–Shannon sampling; often 2.5–3 pixels per resolution unit are helpful, and for some digital image analysis procedures, additional magnification beyond this (‘‘empty magnification’’) may be helpful.

2.6. Other hardware considerations For doing time-lapse measurements of yeast cells, a few additional components will be helpful. Most likely, you will want some kind of temperature control system so that your cells can be grown at a constant temperature. Temperature control can either be achieved by enclosing the entire microscope in a thermostatted plexiglass box, or by using a stage top heater combined with an objective heater. While enclosing the microscope in a temperature controlled box makes access to the microscope and the sample somewhat cumbersome, it does maintain a very stable environment around the entire microscope (typically stable to within 0.1  C). This temperature stability helps minimize focus drift due to thermal fluctuations and makes switching samples and objectives simple. Stage top incubators are much smaller and cheaper and do not encumber the microscope at all. However, for use with oil-immersion objectives, they must be paired with an objective heater as otherwise the objective acts as a heat sink and will chill the sample. As use of an objective heater makes changing objectives difficult, and a stage top incubator will typically only hold a single size of sample dish, this approach adds its own set of difficulties. Another addition which is desirable for time-lapse data acquisition is a motorized X–Y stage. This greatly increases the number of cells that can be recorded in time-lapse by recording multiple fields of view in succession. As typically it will only take 10–15 s to move to a new field of view, autofocus with a hardware autofocus device, and acquire images, a motorized stage will allow tens of fields to be collected during the typical 5 min interval of a time-lapse movie. For cases where fast Z-stack acquisition is required, a piezoelectric Z-stage is a necessary addition. Addition of a piezoelectric Z-stage to either a motorized or a manual stage allows Z-stack acquisition at up to 30 fps if your camera is fast enough and your exposure time is short enough. To minimize vibration and sample drift, you will want to place the entire system on a vibration isolation table.

594

Kurt Thorn

2.7. Software As the spinning-disk scanhead is a relatively passive device (you can set the rotation speed and internal filters), it can be controlled by a large number of software packages. Therefore, the software package you choose to control your microscope will depend more on the other components in your system, particularly the microscope body. Most microscope manufacturers now have their own software packages for controlling their microscopes, which should also be capable of controlling the other components in their systems. In addition, there are a number of third-party software packages that can control most instrumentation. Metamorph, from Molecular Devices, has been a standard third-party software package for many years, and controls a large variety of hardware. More recently, a free, open-source, microscope control package called Micro-Manager has been released. This is an appealing choice for labs looking to save money or who wish to customize their software. Microscope control software will usually include image analysis tools as well and so is generally a good starting point for data analysis.

2.8. System integration In most cases, it does not make sense to purchase all the items described here individually. You will want to work with a system integrator who will sell you the scanhead, lasers and optics, and additional microscope components you need and will assemble the system and get it working. You may need to purchase the microscope stand separately and then add the spinning-disk confocal from a separate vendor. Two system integrators we have had good luck with are Solamere Technologies and Andor Technology.

3. Sample Preparation 3.1. Fluorescent tagging and choice of fluorescent protein One of the major strengths of budding yeast as a cell biological model organism is the ease with which genes can be tagged with fluorescent proteins. Homologous recombination is extremely efficient in budding yeast allowing a fluorescent protein sequence to be integrated nearly anywhere in the genome using a 40-bp sequence to provide targeting. This enables tagging of a specific gene by amplification of a template with PCR primers containing targeting sequences for the gene to be tagged. Providing methods for performing gene tagging is beyond the scope of this review, but protocols are widely available (Amberg et al., 2005; Gauss et al., 2005; Knop et al., 1999; Longtine et al., 1998; Petracek and Longtine, 2002). Because of

Spinning-Disk Confocal Microscopy of Yeast

595

the ease of fluorescent protein tagging in yeast, these are probably the most common fluorescent probes used, although other probes can be used as well (Giepmans et al., 2006). A wide variety of fluorescent proteins (FPs) have been discovered and engineered recently. While a comprehensive discussion of available fluorescent proteins and their properties is not possible given space constraints, several excellent reviews are available (Shaner et al., 2005, 2007; Straight, 2007; Zacharias and Tsien, 2006). Here, I focus on FPs readily available in Saccharomyces cerevisiae tagging vectors and newer proteins with desirable properties (Table 23.1). Tagging vectors are readily available for CFP, GFP, and YFP, and vectors are also available for mCherry, mTFP1, and Sapphire; unpublished vectors also exist for other fluorescent proteins. When choosing a fluorescent protein for spinning-disk confocal imaging of yeast, there are several factors to take into account. Most important is the detectability of the protein—how bright will a tagged protein be relative to the background fluorescence of the cell? Additional factors are the availability of a laser line well matched to the excitation spectrum of the fluorescent protein, the photostability of the fluorescent protein, and whether or not the fluorescent protein perturbs the function of the protein to which it is fused. Detectability of a fluorescent protein is a function of both the brightness of the tagged protein and the autofluorescence of the cell. The intrinsic brightness of the FP plays a major role as, if all other factors are held constant, the brighter the protein, the more detectable it will be. This is easily calculated as the product of the extinction coefficient, which measures how efficiently the fluorophore absorbs light, and quantum yield, which measures how many absorbed photons are reemitted as fluorescence, of the FP; values are given in Table 23.1. Additionally, the maturation rate of the FP will directly affect the brightness of the tagged protein, as a protein with a slow maturation rate will be degraded and diluted by cell growth faster than it matures, leading to a majority of the protein being nonfluorescent. While maturation rates are not known for all FPs, and are frequently measured in vitro or in E. coli, conditions that may not be relevant to yeast expression, the proteins listed in Table 23.1 generally have reasonably fast maturation rates (30 min or less). Similarly, translation efficiency will influence the brightness of a fusion protein; I have found that codon optimization of the FP can give a twofold increase in brightness (Sheff and Thorn, 2004), although mRNA secondary structure may play a larger role (Kudla et al., 2009). The other component of detectability is cellular autofluorescence; the brighter the cellular autofluorescence, the brighter a tagged protein will have to be detected above this background. Careful choice of yeast strain and growth conditions can reduce autofluorescence (see below), but it cannot be completely eliminated. Much of the autofluorescence in yeast is due to flavins, which absorb broadly in the violet–blue range (400–500 nm) and emit in the green (530 nm). Yeast autofluorescence is therefore

Table 23.1 Fluorescent proteins Protein

References

lex

lem

e (M 1 cm 1)

QY

Brightnessa

Tagging vectors

TagBFP Sapphire T-Sapphire

402 399 399

457 511 511

52,000 29,000 44,000

0.63 0.64 0.6

32.8 18.6 26.4

Sheff and Thorn (2004)

ECFP

Subach et al. (2008) Cubitt et al. (1999) Zapata-Hommer and Griesbeck (2003) Tsien (1998)

433

475

32,500

0.4

13.0

mTFP1 EGFP

Ai et al. (2006) Tsien (1998)

462 488

492 507

64,000 56,000

0.85 0.6

54.0 33.6

Citrine

Griesbeck et al. (2001) Nagai et al. (2002) Tsutsui et al. (2008) Shaner et al. (2004) Merzlyak et al. (2007) Wang et al. (2004) Shcherbo et al. (2009)

516

529

83,400

0.76

58.5

515 551 587 555 590 588

529 563 610 584 649 633

92,200 105,000 72,000 100,000 41,000 62,500

0.57 0.61 0.22 0.48 0.1 0.4

52.5 64.0 15.8 49.0 4.1 25.0

Venus mKOk mCherry TagRFP mPlum mKate2

An expanded table is available online at http://thornlab.org/gfps.htm. a Product of e and QY, divided by 1000.

Sheff and Thorn (2004), Janke et al. (2004), Hailey et al. (2002) Deng et al. (2009) Sheff and Thorn (2004), Janke et al. (2004), Deng et al. (2009), Longtine et al. (1998), Gauss et al. (2005), Wach et al. (1997) Sheff and Thorn (2004), Deng et al. (2009) Sheff and Thorn (2004) Deng et al. (2009)

Spinning-Disk Confocal Microscopy of Yeast

597

strongly wavelength dependent, being much less intense for longer wavelength imaging than shorter wavelength imaging. Because of this, orange and red fluorescent proteins are more detectable than would be expected given their intrinsic brightness. The photostability of the fluorescent protein is important when doing time-lapse imaging; more photostable proteins will bleach more slowly and so will allow more frames to be acquired before the signal drops to an unacceptable level. For acquiring single images, photobleaching rate is relatively unimportant. Photobleaching rates have been measured for many of the proteins suggested here and are available in the original publications and in several reviews (Shaner et al., 2005, 2007; Zacharias and Tsien, 2006). Finally, of critical importance is whether or not fusion of a fluorescent protein to your protein of interest will perturb its function. This will of course depend on the protein being tagged and on where it is tagged, but it seems that GFP fusions to the C-terminus of proteins in yeast is generally well tolerated, as 87% of essential yeast genes could be successfully tagged with GFP in a systematic tagging effort (Howson et al., 2005; Huh et al., 2003). In my experience, other GFP variants (e.g., CFP and YFP) are equally well tolerated, and mCherry is well tolerated also. Many of the newer fluorescent proteins mentioned have not been extensively studied in the context of fusion proteins, and so little information is available about how they are tolerated. For general purpose imaging, tagging with GFP is a good place to start. It is readily available in a monomeric form, reasonably bright, and generally well behaved. For imaging a second color, mCherry has minimal cross talk with GFP and is also generally well behaved. For multicolor imaging, CFP/ YFP/mCherry is a good combination, and Sapphire can be added with the introduction of a small amount of cross talk (Sheff and Thorn, 2004). TagBFP is another possibility for a fourth color, and it may be possible to multiplex mKOk for a fifth color with some small amount of cross talk. For imaging low-abundance proteins, it may be worth exploring less common options to maximize signal-to-noise ratio. In general, moving to longer wavelengths is advantageous as yeast autofluorescence is less intense at longer wavelengths. For proteins that are rapidly turned over, using a fastfolding protein such as Venus may help (Yu et al., 2006). For difficult imaging cases, I would expect that it may be necessary to try several proteins before identifying an optimal tag. Finally, imaging in diploid cells may be preferred to imaging in haploid cells as diploid cells are larger.

3.2. Minimizing autofluorescence Yeast cells are somewhat autofluorescent, and the commonly used ade2 strains accumulate a highly fluorescent red pigment (Ishiguro, 1989; Stotz and Linder, 1990). If possible, it is best to avoid ade2 strains, although accumulation of this pigment can be avoided by supplementing the growth

598

Kurt Thorn

medium with 20 mg/ml adenine. This supplementation can also be beneficial for strains that are ADEþ. Cells tend to be less fluorescent in early log phase (OD600  0.2). Yeast media can also be autofluorescent. Rich media such as YPD are highly autofluorescent and should be avoided when imaging. Cells grown in rich media can be washed into a less fluorescent media or buffer prior to imaging; however, these fluorescent media seem to increase the autofluorescence of cells slightly even after washing. In synthetic (minimal) media, the major sources of autofluorescence are riboflavin and folic acid, and omitting these components eliminates the media autofluorescence if the medium is made from scratch. Most S. cerevisiae strains can synthesize both riboflavin and folic acid, so eliminating these vitamins from the medium does not appear to have drawbacks. Commercial yeast nitrogen base is often fluorescent even when it is lacking these two vitamins, so I recommend testing its fluorescence before use or making your own from scratch (Sheff and Thorn, 2004). Commercial CSM supplements do not seem to be autofluorescent and so can be safely used. Growing cells in this low-fluorescence medium allows direct imaging of cultures without washing, and substantially reduces backgrounds for cells grown on agarose pads or in a perfusion system.

3.3. Mounting The simplest way to prepare yeast for imaging for short periods of time (30–60 min) is by immobilizing them on Concanavalin A coated coverslips. Concanavalin A binds to sugars in the yeast cell wall and will stick the yeast tightly to the coverslip so that they will not move during imaging. Concanvalin A coated coverslips can easily be prepared as follows: 1. Prepare Concanavalin A solution by dissolving Concanavalin A (Sigma #L 7647) in distilled water to 0.5 mg/ml. Refrigerate. 2. Rack 22 mm #1.5 coverslips (Racks: Electron Microscopy Sciences #72241-01). 3. Soak coverslips overnight in 1 M NaOH with gentle shaking. Sterile filter NaOH before use to remove dust. 4. Pour off NaOH and save for reuse. Wash coverslips 3 with distilled water. 5. Add Concanavalin A solution and soak for 20 min with gentle shaking. 6. Pour off Concanavalin A solution and save for reuse. 7. [Optional] Rinse coverslips once with distilled water. 8. [Optional] Spin coverslips dry on microplate carriers in Sorvall RT7. Place racks on a piece of paper towel to catch liquid and spin 1 min at 700 rpm. 9. Place racks in hood until absolutely dry. Store in coverslip box at RT. Coverslips should be good for at least 1 month.

Spinning-Disk Confocal Microscopy of Yeast

599

To use the coverslips, simply place a 6–8 ml drop of yeast cell suspension on a slide, and drop a coated coverslip on it. The cells should stick to the coverslip within a few minutes. The coverslip can be sealed with petroleum jelly or VALAP (Vaseline:Lanolin:Paraffin, 1:1:1) for longer term imaging, or left unsealed for short-term studies (10 min). If the cells are grown in low-fluorescence media, they can be directly placed on coverslips for imaging; if they are grown in more fluorescent media, particularly in YPD or other rich media, you will substantially reduce the background fluorescence by first washing them into PBS or some other nonfluorescent buffer or medium. Pelleting and resuspending cells can also be used to concentrate the cells if the cell density in the initial culture is not high enough to get a sufficient number of cells in the field of view. For longer term imaging (up to 4–6 h), cells can be grown on agarose pads containing low-fluorescent yeast medium. These can be prepared by dissolving 1.2% agarose in low-fluorescence SC þ carbon source. The agarose solution is then cast in a 1-mm slab in a device used for casting polyacrylamide gels. If kept in a sealed container with moist paper towels, such a gel can be kept for roughly a week. Pieces of agarose ( 15  15 mm) are then cut out with a sterilized razor blade and placed on a slide, a drop of yeast suspension is placed on the agarose block, and a coverslip is placed on top. The gap between the coverslip and slide can then be filled with petroleum jelly to prevent evaporation. This is easily done by filling a syringe with petroleum jelly and injecting it through a large gauge needle into the space between the coverslip and slide. Alternatively, these pads can be sealed with VALAP. Agarose pads can also be made by filling depression slides with agarose, or sandwiching a drop of agarose between two slides, but I find the casting method described here to be the easiest. For even longer term imaging (overnight or longer), it is probably best to use a microfluidic device to provide constant nutrient flow to the cells and remove waste products. We have had good luck with the CellASIC ONIXTM system (www.cellasic.com). This system consists of a control device paired with a special microfluidic plate which has the same footprint as an ordinary 96-well plate. The plate contains viewing areas where cells can be trapped and immobilized for long-term imaging while being continuously perfused with solution. The cells are trapped and held in place by being loaded under pressure (6–8 psi) into a viewing area which traps the cells under an elastomeric membrane 4.2 mm above the coverslip. The downward pressure applied by the ceiling membrane holds the cells in place indefinitely. Media can be perfused from one of two wells, allowing rapid media switching. Cells have been kept in this device for up to 3 days and continued to divide (Lee et al., 2008).

600

Kurt Thorn

ACKNOWLEDGMENTS Thanks to S. Rafelski, C. Mrejen, and G. Peeters for helpful discussions. Thanks to the many users of the Nikon Imaging Center spinning-disk confocal for discussions and insight into how to optimize spinning-disk confocal microscopy for specific applications. The images in Fig. 23.2 were acquired in the Nikon Imaging Center at UCSF/QB3.

REFERENCES Ai, H. W., et al. (2006). Directed evolution of a monomeric, bright and photostable version of Clavularia cyan fluorescent protein: Structural characterization and applications in fluorescence imaging. Biochem. J. 400(3), 531–540. Amberg, D. C., Burke, D., and Strathern, J. N. Cold Spring Harbor Laboratory (2005). Methods in yeast genetics: A Cold Spring Harbor Laboratory course manual. 2005 edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Art, J. (2006). Photon detectors for confocal microscopy. In ‘‘Handbook of Biological Confocal Microscopy,’’ ( J. Pawley, ed.), 3rd edn. Springer ScienceþBusiness Media, New York, NY. Booth, M. J. (2007). Adaptive optics in microscopy. Phil. Trans. A Math. Phys. Eng. Sci. 365, 2829–2843. Cubitt, A. B., Woollenweber, L. A., and Heim, R. (1999). Understanding structurefunction relationships in the Aequorea victoria green fluorescent protein. Methods Cell Biol. 58, 19–30. Davis, T. N. (2004). Protein localization in proteomics. Curr. Opin. Chem. Biol. 8, 49–53. Deng, C., Xiong, X., and Krutchinsky, A. N. (2009). Unifying fluorescence microscopy and mass spectrometry for studying protein complexes in cells. Mol. Cell Proteomics. 8, 1413–1423. Gauss, R., Trautwein, M., Sommer, T., and Spang, A. (2005). New modules for the repeated internal and N-terminal epitope tagging of genes in Saccharomyces cerevisiae. Yeast 22, 1–12. Giepmans, B. N., Adams, S. R., Ellisman, M. H., and Tsien, R. Y. (2006). The fluorescent toolbox for assessing protein location and function. Science 312, 217–224. Griesbeck, O., et al. (2001). Reducing the environmental sensitivity of yellow fluorescent protein. Mechanism and applications. J. Biol. Chem. 276(31), 29188–29194. Hailey, D. W., Davis, T. N., and Muller, E. G. (2002). Fluorescence resonance energy transfer using color variants of green fluorescent protein. Methods Enzymol. 351, 34–49. Howson, R., Huh, W. K., Ghaemmaghami, S., Falvo, J. V., Bower, K., Belle, A., Dephoure, N., Wykoff, D. D., Weissman, J. S., and O’Shea, E. K. (2005). Construction, verification and experimental use of two epitope-tagged collections of budding yeast strains. Comp. Funct. Genomics 6, 2–16. Huh, W. K., Falvo, J. V., Gerke, L. C., Carroll, A. S., Howson, R. W., Weissman, J. S., and O’Shea, E. K. (2003). Global analysis of protein localization in budding yeast. Nature 425, 686–691. Inoue, S. (2006). Foundations of confocal scanned imaging in light microscopy. In ‘‘Handbook of Biological Confocal Microscopy,’’ ( J. Pawley, ed.), 3rd edn. Springer ScienceþBusiness Media, New York, NY. Ishiguro, J. (1989). An abnormal cell division cycle in an AIR carboxylase-deficient mutant of the fission yeast Schizosaccharomyces pombe. Curr. Genet. 15, 71–74. Janesick, J. R. (2001). Scientific Charge-Coupled Devices. SPIE Press, Bellingham, Washington.

Spinning-Disk Confocal Microscopy of Yeast

601

Janke, C., et al. (2004). A versatile toolbox for PCR-based tagging of yeast genes: New fluorescent proteins, more markers and promoter substitution cassettes. Yeast 21(11), 947–962. Keller, H. E. (2006). Objective lenses for confocal microscopy. In ‘‘Handbook of Biological Confocal Microscopy,’’ ( J. Pawley, ed.), 3rd edn. Springer ScienceþBusiness Media, New York, NY. Knop, M., Siegers, K., Pereira, G., Zachariae, W., Winsor, B., Nasmyth, K., and Schiebel, E. (1999). Epitope tagging of yeast genes using a PCR-based strategy: More tags and improved practical routines. Yeast 15, 963–972. Kohlwein, S. D. (2000). The beauty of the yeast: Live cell microscopy at the limits of optical resolution. Microsc. Res. Tech. 51, 511–529. Kudla, G., Murray, A. W., Tollervey, D., and Plotkin, J. B. (2009). Coding-sequence determinants of gene expression in Escherichia coli. Science 324, 255–258. Lee, P. J., Helman, N. C., Lim, W. A., and Hung, P. J. (2008). A microfluidic system for dynamic yeast cell imaging. Biotechniques 44, 91–95. Longtine, M. S., McKenzie, A. 3rd, Demarini, D. J., Shah, N. G., Wach, A., Brachat, A., Philippsen, P., and Pringle, J. R. (1998). Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14, 953–961. McAllister, R. G., Sisan, D. R., and Urbach, J. S. (2008). Design and optimization of a highspeed, high-sensitivity, spinning disk confocal microscopy system. J. Biomed. Opt. 13, 054058. Merzlyak, E. M., et al. (2007). Bright monomeric red fluorescent protein with an extended fluorescence lifetime. Nat. Methods 4(7), 555–557. Murray, J. M., Appleton, P. L., Swedlow, J. R., and Waters, J. C. (2007). Evaluating performance in three-dimensional fluorescence microscopy. J. Microsc. 228, 390–405. Nagai, T., et al. (2002). A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat. Biotechnol. 20(1), 87–90. Pawley, J. (2006). Points, pixels, and gray levels, digitizing image data. In ‘‘Handbook of Biological Confocal Microscopy,’’ ( J. Pawley, ed.), 3rd edn. Springer ScienceþBusiness Media, New York, NY. Petracek, M. E., and Longtine, M. S. (2002). PCR-based engineering of yeast genome. Methods Enzymol. 350, 445–469. Rines, D. R., He, X., and Sorger, P. K. (2002). Quantitative microscopy of green fluorescent protein-labeled yeast. Methods Enzymol. 351, 16–34. Shaner, N. C., et al. (2004). Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein. Nat. Biotechnol. 22(12), 1567–1572. Shaner, N. C., Steinbach, P. A., and Tsien, R. Y. (2005). A guide to choosing fluorescent proteins. Nat. Methods 2, 905–909. Shaner, N. C., Patterson, G. H., and Davidson, M. W. (2007). Advances in fluorescent protein technology. J. Cell Sci. 120, 4247–4260. Shaw, P. J. (2006). Comparison of widefield/devonvolution and confocal microscopy for three-dimensional imaging. In ‘‘Handbook of Biological Confocal Microscopy,’’ ( J. Pawley, ed.), 3rd edn. Springer ScienceþBusiness Media, New York, NY. Shcherbo, D., et al. (2009). Far-red fluorescent tags for protein imaging in living tissues. Biochem. J. 418(3), 567–574. Sheff, M. A., and Thorn, K. S. (2004). Optimized cassettes for fluorescent protein tagging in Saccharomyces cerevisiae. Yeast 21, 661–670. Stotz, A., and Linder, P. (1990). The ADE2 gene from Saccharomyces cerevisiae: Sequence and new vectors. Gene 95, 91–98. Straight, A. F. (2007). Fluorescent protein applications in microscopy. Methods Cell Biol. 81, 93–113.

602

Kurt Thorn

Subach, O. M., et al. (2008). Conversion of red fluorescent protein into a bright blue probe. Chem. Biol. 15(10), 1116–1124. Toomre, D., and Pawley, J. B. (2006). Disk-scanning confocal microscopy. In ‘‘Handbook of Biological Confocal Microscopy,’’ ( J. Pawley, ed.), 3rd edn. Springer ScienceþBusiness Media, New York, NY. Tsien, R. Y. (1998). The green fluorescent protein. Annu. Rev. Biochem. 67, 509–544. Tsutsui, H., et al. (2008). Improving membrane voltage measurements using FRET with new fluorescent proteins. Nat. Methods 5(8), 683–685. Wach, A., et al. (1997). Heterologous HIS3 marker and GFP reporter modules for PCRtargeting in Saccharomyces cerevisiae. Yeast 13(11), 1065–1075. Wang, L., et al. (2004). Evolution of new nonantibody proteins via iterative somatic hypermutation. Proc. Natl. Acad. Sci. USA 101(48), 16745–16749. Yu, J., Xiao, J., Ren, X., Lao, K., and Xie, X. S. (2006). Probing gene expression in live cells, one protein molecule at a time. Science 311, 1600–1603. Zacharias, D. A., and Tsien, R. Y. (2006). Molecular biology and mutation of green fluorescent protein. Methods Biochem. Anal. 47, 83–120. Zapata-Hommer, O., and Griesbeck, O. (2003). Efficiently folding and circularly permuted variants of the Sapphire mutant of GFP. BMC Biotechnol. 3, 5.